A clean blood draw and gentle handling of specimens are required to avoid spontaneous platelet activation. If agonists are being added during processing to study cell reactivity, platelet aggregation could occur and interfere with analysis. It is important, therefore, that the samples be left undisturbed during any activation steps, followed immediately by fixation. Even in the absence of added agonist, LPA will form relatively quickly once blood is drawn into anticoagulant; therefore, the time to processing should be kept to a minimum (<20 min) to avoid artifactual LPA formation. Large platelet-platelet aggregates can exhibit high light scatter properties, and can interfere with analysis. These can generally be gated out, however, by taking advantage of their high platelet-specific and low leukocyte-specific fluorescence. EDTA-anticoagulated whole blood should never be used for LPA analysis, as the EDTA can cause in vitro dissociation of platelets from leukocytes.
To minimize delays, all tubes should be labeled and reagents prepared prior to obtaining the blood sample. Anticoagulants such as CTAD (citrate theophylline adenosine dipyridimole) or added platelet activation inhibitors, such as prostacyclin or apyrase, can minimize formation of spontaneous LPAs, but these methods will alter agonist responsiveness and may even result in dissaggregation of existing LPA. Samples should never be exposed to temperatures <15°C, as rewarming will result in platelet degranulation and LPA formation.
The precise percent LPAs obtained using the Basic Protocol may differ depending on the antibodies chosen as leukocyte- and platelet-specific markers. CD14 was chosen here since it does not appear to play a role in leukocyte-platelet aggregation. The influence of leukocyte-specific antibodies on LPA formation can be determined by comparing LPAs found by gating on the leukocyte-specific antibody signal versus side light scatter to those found by gating on forward versus side light scatter in the absence of the leukocyte antibody. The monocyte population in particular will be different (lower purity) but differences due to the presence of antibody can be estimated. The platelet-specific antibody can also influence LPA development. For example, antibodies directed toward the Mac-1 (CD11b/18) binding region of platelet CD42b should be avoided as this interaction is important in the stabilization of LPAs (Simon et al., 2000). Blocking antibodies for P-selectin (CD62P) or its counter-receptor PSGL-1 (CD162) are useful for preparing baseline control samples as they are very effective in blocking formation of LPAs. These blocking agents will not dissociate stable LPAs, however, so these can not be considered true negative controls. Significant differences in the baseline LPA level in the presence or absence of blocking antibody may indicate an activating effect by the chosen leukocyte or platelet antibody.
The pre-fixation method described in the Alternate Protocol is less susceptible to antibody influences, but sample fixation and washing can also modify the measurement of LPA. Fixation can reduce or eliminate binding of some antibodies. As a general rule, anti-platelet antibodies, which label with greater intensity, will result in an increase in the apparent number of LPAs. This is particularly noticeable in the baseline sample, in which few platelets are bound per LPA. The pre-fixation technique requires washing to remove the fixative and concentrate the sample for labeling. Care must also be taken during wash steps since even fixed CD62P-positive platelets can bind leukocytes. The low G force described in the Alternate Protocol does not pellet most free platelets, so this effect is minimized in the method described.
Monocytes are the time-limiting cell type during data acquisition; 2 to 7 min are needed to acquire 1000 to 2000 monocyte events. Samples should be run on a low to medium flow rate, depending on dilution. It is tempting to concentrate samples or increase flow rates; however, leukocyte-platelet coincident events (particularly monocyte-platelet events) are a distinct problem due to the large number of platelets in the unwashed whole blood samples described in this method. To determine the extent to which coincidence is contributing to increased leukocyte-platelet events, accumulate 1000 monocyte events at high, medium, and low flow settings with the same sample. Typically, the baseline (no added agonist) percent of monocytes that are platelet positive will be considerably increased for samples run at high flow rates. In some circumstances, a low flow rate should be used, particularly if the platelet count in the sample is abnormally high, but at the final blood dilution described in this protocol (1:44) medium flow is acceptable for most samples. Sheath pressures at medium and low flow settings determine absolute flow rate and core stream diameter and will differ from one instrument to another. Therefore, each laboratory should independently determine the optimal flow rate for a given instrument. Samples should be run as soon as possible after fixation, although LPAs labeled and subsequently fixed are quite stable up to 72 hours when stored at 4°C.
LPA results are usually expressed as the percentage of the gated leukocyte subset that stains positively for the platelet-specific antibody. In addition to neutrophils and monocytes, which constitute the majority of leukocytes in circulating LPAs, other leukocyte subsets such as eosinophils, basophils, and NK cells are capable of binding platelets (Bruijne-Admiraal et al., 1992).
Approximately 4% to 14% of monocytes from healthy normal blood donors (citrate-an-ticoagulated whole blood, no agonist added) will be positive for platelet markers (i.e., have attached platelets), while ~2% to 12% of neu-trophils in the same sample will be positive for platelet markers. Even a carefully drawn sample from a patient with a recent thrombotic event, such as an acute myocardial infarction, can have as many as 80% circulating monocyte-platelet aggregates. A normal donor sample stimulated in vitro with 20 |M TRAP will become nearly 100% monocyte-platelet positive with neutrophils becoming 80% to 95% platelet positive in the same sample.
The mean fluorescence of a given LPA type generally reflects the relative number of platelets bound to that leukocyte subtype. However, it is difficult to translate this into the exact number of platelets per LPA due to several factors. Normal platelets and platelet-derived microparticles vary widely in size and expression of surface antigens, and thus the amount of fluorescence for a single bound platelet varies widely. In addition, the membrane-binding domain between the leukocyte and platelet can cause loss of available surface for staining, and the presence of a large number of fluorophore molecules in proximity can result in quenching of emitted fluorescence. For these reasons, relative LPA fluorescence should be considered semi-quantitative.
A significant portion of the time commitment for LPA study is devoted to the necessary reagent optimization and generation of a normal range for a given assay configuration. Sample stability must also be confirmed in clinical studies in which immediate access to a flow cytometer is not always practical. Once a procedure has been established, the assay should be set up in anticipation of obtaining the blood sample (~20 to 30 min). Whole blood labeling with or without activation should take <20 min at 22° to 37°C. Subsequent fixation can be done in 10 min. The most time-consuming hands-on portion of LPA analysis may be data acquisition, which can take 2 to 7 min per sample to acquire 1000 to 2000 monocytes.
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